857
Neuroscience
J Physiol 593.4 (2015) pp 857–870
Proton transfer unlocks inactivation in cyclic
nucleotide-gated A1 channels
Arin Marchesi, Manuel Arcangeletti, Monica Mazzolini and Vincent Torre
Neurobiology Sector, International School for Advanced Studies (SISSA), Trieste, Italy
Key points
r Desensitization and inactivation provide a form of short-term memory controlling the firing
patterns of excitable cells and adaptation in sensory systems.
r Unlike many of their cousin K+ channels, cyclic nucleotide-gated (CNG) channels are thought
The Journal of Physiology
not to desensitize or inactivate.
r Here we report that CNG channels do inactivate and that inactivation is controlled by
extracellular protons.
r Titration of a glutamate residue within the selectivity filter destabilizes the pore architecture,
r
which collapses towards a non-conductive, inactivated state in a process reminiscent of the
usual C-type inactivation observed in many K+ channels.
These results indicate that inactivation in CNG channels represents a regulatory mechanism
that has been neglected thus far, with possible implications in several physiological processes
ranging from signal transduction to growth cone navigation.
Abstract Ion channels control ionic fluxes across biological membranes by residing in any of
three functionally distinct states: deactivated (closed), activated (open) or inactivated (closed).
Unlike many of their cousin K+ channels, cyclic nucleotide-gated (CNG) channels do not
desensitize or inactivate. Using patch recording techniques, we show that when extracellular
pH (pHo ) is decreased from 7.4 to 6 or lower, wild-type CNGA1 channels inactivate in a
voltage-dependent manner. pHo titration experiments show that at pHo < 7 the I–V relationships
are outwardly rectifying and that inactivation is coupled to current rectification. Single-channel
recordings indicate that a fast mechanism of proton blockage underlines current rectification while
inactivation arises from conformational changes downstream from protonation. Furthermore,
mutagenesis and ionic substitution experiments highlight the role of the selectivity filter in current
decline, suggesting analogies with the C-type inactivation observed in K+ channels. Analysis with
Markovian models indicates that the non-independent binding of two protons within the transmembrane electrical field explains both the voltage-dependent blockage and the inactivation.
Low pH, by inhibiting the CNGA1 channels in a state-dependent manner, may represent an
unrecognized endogenous signal regulating CNG physiological functions in diverse tissues.
(Received 10 September 2014; accepted after revision 28 November 2014; first published online 4 December 2014)
Corresponding author V. Torre: Neurobiology Sector, International School for Advanced Studies (SISSA), Via Bonomea
no. 265-34136, Trieste, Italy. Email: [email protected]
Abbreviations BS, binding site; CN, cyclic nucleotide; CNG, cyclic nucleotide-gated; VGIC, voltage-gated ion channel.
Introduction
Cyclic nucleotide-gated (CNG) channels conduct a
Ca2+ -permeable non-selective cation current mediating
A. Marchesi and M. Arcangeletti contributed equally to this work.
signal transduction in photoreceptor and olfactory
sensory neurons (Hille, 1992; Kaupp & Seifert, 2002;
Craven & Zagotta, 2006). Although CNG channels belong
to the superfamily of voltage-gated ion channels (VGICs),
they open in response to binding of cyclic nucleotides
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
This is an open access article under the terms of the Creative Commons Attribution NonCommercial License, which permits
use, distribution and reproduction in any medium, provided the original work is properly cited and is not used for commercial
purposes.
[The copyright line for this article was changed on 17th June 2016 after original online publication]
DOI: 10.1113/jphysiol.2014.284216
858
A. Marchesi and others
(CNs) and are only barely modulated by membrane
voltage (Hille, 1992; Kaupp & Seifert, 2002; Yu et al.
2005; Craven & Zagotta, 2006; Mazzolini et al. 2010). CNG
channel activity is also regulated by a variety of molecules
and ions, including protons. Changes in extracellular
hydrogen ion concentration (pHo ) occur in a variety of
physiological and pathophysiological conditions, such as
neuronal activity, ischaemia and inflammation (Kellum
et al. 2004; Isaev et al. 2008; Magnotta et al. 2012). In
the retina, pHo follows a circadian rhythm (Dmitriev
& Mangel, 2001), and pH changes could play a role
in adaptation of the retinal response to different light
intensities. Moreover, acidosis has been associated with
changes in cell excitability in vascular tissues as well as in
the CNS (Tolner et al. 2011; Pavlov et al. 2013), in which
CNG channels are widely expressed (Zufall et al. 1997;
Kaupp & Seifert, 2002; Leung et al. 2010; Lopez-Jimenez
et al. 2012). Therefore, determining the effect of pHo on
CNG channel gating has great clinical and physiological
significance.
It is well established that extracellular protons inhibit
CNG currents, although conflicting mechanisms have
been proposed (Root & MacKinnon, 1994; Rho & Park,
2013; Morrill & MacKinnon, 1999; Martı́nez-François
et al. 2010). For instance, the outward rectification
observed at low pHo has been attributed to a
voltage-dependent proton blockage (Rho & Park, 2013)
and more recently to an enhancement of the wild-type
(WT) channel inherent mild voltage-dependent gating
(Martı́nez-François et al. 2010). Here we show that
when pHo is decreased from 7.4 to 6 or lower, WT
CNGA1 channels inactivate or desensitize. The term
‘desensitization’ usually refers to a mechanism of activity
attenuation following sustained exposure to a chemical
messenger, such as a neurotransmitter in ligand-gated
channels, whereas inactivation is commonly used for
VGICs. In the present investigation, mutagenesis and
ionic substitution experiments indicated that the current
decline is reminiscent of the usual C-type inactivation
observed in Kv channels and consequently we refer
to the observed loss of conduction as inactivation. A
simple kinetic model in which two protons bind within
the selectivity filter provides a conceptual framework
to rationalize both voltage-dependent proton block
and inactivation. Inactivation thus represents a novel
regulatory mechanism of CNG channels that has been
neglected.
Methods
Ethical approval
All studies were approved by the SISSA’s Ethics Committee
according to the Italian and European guidelines for
animal care (d.l. 116/92; 86/609/C.E.). Oocytes were
J Physiol 593.4
harvested from female Xenopus laevis frogs using an
aseptic technique. All X. laevis surgeries were performed
under general anaesthesia, obtained by immersion in a
0.2% solution of tricaine methane sulfonate (MS-222)
adjusted to pH 7.4 for 15–20 min. Depth of anaesthesia
was assessed by loss of the righting reflex and loss of
withdrawal reflex to a toe pinch. After surgery animals
were singly housed for 48 h. Frogs were monitored daily
for 1 week post-operatively to ensure the absence of
any surgery-related stress. Post-operative analgesics were
not routinely used. Considering the simplicity of the
procedure, the lack of complications, the effectiveness
of anaesthetic regimens and reductions in the number
of animals likely to occur compared to the number that
would be required if only one surgery were permitted,
multiple surgeries on a single animal were performed.
Individual donors were used up to five times, conditional
upon the relative health of an individual animal. Recovery
time between oocyte collection from the same animal
was maximized by rotation of the frogs being used.
A minimum recovery period of 1 month was ensured
between ovarian lobe resection from the same animal to
avoid distress. Evidence of surgery-related stress resulted
in an extended rest period based on recommendations
from the veterinary staff. After the fifth terminal surgery
frogs were humanely killed through anaesthesia overdose
via 2 h of immersion in a 5 g l−1 MS-222 solution adjusted
to pH 7.4.
Molecular biology
The CNGA1 channel from bovine rods consisting of 690
amino acids was used (Kaupp et al. 1989). Selected residues
were replaced as described (Becchetti et al. 1999) using the
Quick Change Site-Directed Mutagenesis kit (Stratagene,
La Jolla, CA, USA). Point mutations were confirmed by
sequencing, using a LI-COR sequencer (4000 l; LI-COR
Biosciences, Lincoln, NE, USA). cDNAs were linearized
and were transcribed to cRNA in vitro using the mMessage
mMachine kit (Ambion, Austin, TX, USA).
Oocyte preparation and chemicals
Mutant channel cRNAs were injected into X. laevis oocytes
(‘Xenopus express’ Ancienne Ecole de Vernassal, Le Bourg
43270, Vernassal, Haute-Loire, France). Oocytes were prepared as described (Nizzari et al. 1993). Injected eggs were
maintained at 18°C in a Barth solution supplemented with
50 μg ml–1 gentamycin sulfate and containing (in mM):
88 NaCl, 1 KCl, 0.82 MgSO4 , 0.33 Ca(NO3 )2 , 0.41 CaCl2 ,
2.4 NaHCO3 and 5 Tris-HCl, pH 7.4 (buffered with
NaOH). During the experiments, oocytes were kept in
a Ringer solution containing (in mM): 110 NaCl, 2.5 KCl,
1 CaCl2 , 1.6 MgCl2 and 10 Hepes, pH 7.4 (buffered with
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
J Physiol 593.4
Proton transfer unlocks inactivation in CNGA1 channels
NaOH). Usual salts and reagents were purchased from
Sigma Chemicals (St Louis, MO, USA).
Recording apparatus
cGMP-gated currents from excised patches (Hamill
et al. 1981) were recorded with a patch-clamp amplifier
(Axopatch 200; Axon Instruments Inc., Foster City, CA,
USA), 2–6 days after RNA injection, at room temperature
(20–24°C). The perfusion system was as described (Sesti
et al. 1995) and allowed a complete solution change in
less than 0.1 s. Macroscopic and single-channel current
recordings were obtained with borosilicate glass pipettes
which had resistances of 2–5 M in symmetrical standard
solution. The standard solution on both sides of the
membrane consisted of (in mM) 110 NaCl, 10 Hepes and
0.2 EDTA (pH 7.4). For solutions buffered at pH lower
than 7, 10 mM MES instead of 10 mM Hepes was used and
for solution at pH 9, 10 mM CHES was used. Solutions
were buffered with tetramethylammonium hydroxide at
the desired pH. When the cation X+ was used as the charge
carrier, NaCl in the standard solution on both sides of the
membrane patch was replaced by an equimolar amount
of the cation X+ . We used Clampex version 10.0 for
data acquisition. Recordings from macroscopic currents
to elicit I–V relationships (Figs 2A, B, G and 4I) and to
perform noise analysis (Fig. 2D) were low-pass filtered at
10 kHz. Current signals were sampled with a 16-bit A/D
converter (Digidata 1440A; Axon Instruments), using a
sampling rate of 50 kHz. All other macroscopic current
recordings were low-pass filtered at 1 kHz and sampled
at 2.5 kHz if not otherwise indicated. Single-channel
recordings were low-pass filtered at 2 kHz and sampled
at 5 kHz.
Data analysis
Inactivation time constants (τ) were obtained from
fitting cGMP activated currents with one-component
exponential functions. Single-channel currents (i) were
estimated from patches containing only one CNGA1
channel and fitting normalized all-point histograms
with two-component Gaussian functions as previously
described (Bucossi et al. 1997). The dependency of the
fraction of unblocked channels (Punblocked /Pmax , Fig. 2I) on
voltage at different pHo was determined from the ratios
of the normalized conductance G/G+200 at the indicated
pHo and at pHo = 9. If not otherwise indicated, data
are presented as mean ± SEM, with n indicating the
number of patches. Statistical significance for parametric
analysis was determined using unpaired two-tailed t-test
or single-variable ANOVA, as indicated. For pairwise
comparisons, a Holm–Sidak test was used as post hoc
test. P < 0.01 or P < 0.05 were considered significant,
859
as indicated. Data analysis and figures were made with
Clampfit version 10.1 (Molecular Devices, Sunnyvale,
CA, USA) and Sigmaplot 12.0 (Systat Software, Chicago,
IL, USA). Kinetic models were generated and evaluated
using MatLab 7.9.0 (MathWorks, Natick, MA, USA). The
voltage-dependent association constant for protons shown
in Fig. 6 was assumed to be of the form:
X (V) = X (0)e zδVF /RT
where X(0) is the association constant at 0 mV, z is the
valence (+1), V is the voltage across the membrane, δ is
the fraction of the membrane voltage at the binding site
(the electrical distance) and F/RT = 25.5 mV at 23°C.
Results
Extracellular protons induce inactivation and
outward rectification
WT CNG channels are known not to inactivate or to
desensitize in the presence of a steady CN concentration.
Rather unexpectedly, when CNGA1 channels were
exposed to a low pHo , they inactivated (Fig. 1A). We
filled the patch pipette with the usual concentration of
110 mM NaCl buffered at pHo 5 and stepped the voltage
command to −50 mV (Fig. 1A). After a few seconds we
added 1 mM cGMP to the medium bathing the cytoplasmic
side of the membrane patch (indicated by solid blue bars
in Fig. 1A) and very quickly the expected cGMP-gated
current appeared (Fig. 1A). This current, however, was not
stable and declined to a much lower level corresponding to
approximately 25% of the current initially observed. After
30–40 s of exposure to cGMP, the CN was removed from
the medium and we monitored the current amplitude by
exposing the membrane patch to brief pulses of cGMP.
The amplitude of the cGMP-activated current recovered
to its original level, provided that cGMP was removed for
at least 60–90 s from the bathing medium.
Inactivation was not observed when pHo was reduced
from 7.4 to 7 (Fig. 1C) or when increased to 8 (Fig. 1B).
When pHo was reduced to 6, an appreciable inactivation
was observed only at negative voltages (Fig. 1D). At
pHo 5, CNG channels inactivate significantly also at
positive voltages (Fig. 1E). At steady state, currents
declined to 72 ± 6 and 28 ± 2% (n = 4) of their
initial values at +60 and −60 mV, respectively. When
inactivation fully developed, after a steady exposure to
cGMP for 1–2 min, the I–V relationships were clearly
outward rectifying at pHo 5 (Fig. 2A), in agreement with
previous observations (Martı́nez-François et al. 2010).
Thus, extracellular acidification not only induces current
rectification (Rho & Park, 2013; Martı́nez-François et al.
2010) but is also responsible for a slow, state-dependent
loss of conduction modulating CNG channel activity.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
860
A. Marchesi and others
and exhibits a slow run-down as observed in inside-out
patches at the same pHo (Fig. 1E). The current decline
is fully reversible when pHo was reverted from 5 to 7.4
(Fig. 1F, right panel).
As inactivation appears to be stronger at negative
potentials (Fig. 1E) and both inactivation and rectification
depend on pHo (Fig. 1B–E and G), the observed outward
rectification (Fig. 2A) could arise from the voltage and
pH dependency of the current decline. However, several
observations suggest that this is not the case. First, while
The effect of extracellular acidification is best seen in
outside-out membrane patches where different proton
concentrations could be assessed during the same
experiment (Fig. 1F). Mg2+ at 20 mM was used (indicated
by solid red bars in Fig. 1F) to block CNG channels –
which are continuously exposed to 1 mM cGMP present
in the patch pipette – and estimate the leak current (Liu
& Siegelbaum, 2000). Upon pHo switch to 5, a current
corresponding to approximately 20% of the current at pHo
7.4 is observed (Fig. 1F). Notably, this current is not stable
A
J Physiol 593.4
pHо 5
0 mV
–50 mV
+cGMP
20 pA
20 s
B
C
pHо 8
D
pHо 7
0.5 nA
0.5 nA
pHо 5
0.5 pA
0.1 nA
F
15 s
0 mV
–50 mV
pHо 5
pHо 7.4
E
pHо 6
+Mg2+
0 pA
0.5 s
0.5 nA
20 s
i
G
ii
H
1
rel|l–60/l+60|
1.0
|l–60/l+60|
ii
i
0.5
peak
l-V
0.0
peak
inactivated
0.1
5
6
7
pHо
8
9
5
6
7
pHо
8
9
Figure 1. Properties of macroscopic currents at
different pHo in WT CNGA1 channel
A, current recording during a voltage step at −50 mV at
pHo 5 (pHi 7.4) in symmetrical 110 mM Na+ solutions.
Currents were evoked by the application of 1 mM cGMP
as indicated by solid blue bars. Brief cGMP pulses were
used to monitor channel recovery from inactivation. A
second prolonged application of cGMP after recovery
produced an almost identical time-dependent current
decline. B–E, current recordings during voltage steps at
+60 and −60 mV evoked by 1 mM cGMP jumps at pHo 8
(B), 7 (C), 6 (D) and 5 (E). F, current recording obtained
from a representative outside-out patch, during a
prolonged voltage step at −50 mV, in symmetrical
110 mM Na+ solutions. The experiment was performed
varying pHo from 7.4 to 5 and back to 7.4 (solid blue line).
CNG channels were continuously activated by 1 mM cGMP
present in the patch pipette. Reference current was
obtained by adding 20 mM Mg2+ to the bath solution as
indicated (solid red lines). Kinetics of current activation at
pHo 7.4 prior to (i) and after inactivation at pHo 5 (ii) are
shown to the right. The areas in the green and red boxes
are expanded along the time scale and superimposed for
comparison. G, dependence of current rectification at
+60 and −60 mV (|I–60 /I+60 |) on pHo determined before
inactivation onset (green circles) and from I–V
relationships such as in Fig. 2A (black circles). H,
dependence of relative current rectification (rel|I–60 /I+60 |)
on pHo before (green circles) and after (red circles) current
inactivation. rel|I–60 /I+60 | was determined from the ratio of
I–60 /I+60 at the indicated pHo and I–60 /I+60 at pHo 9.
Green and red arrows in B–E refer to the currents prior to
and after inactivation development. Dashed black lines
indicate the 0 current level.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
J Physiol 593.4
Proton transfer unlocks inactivation in CNGA1 channels
861
rectification measured at +60 and −60 mV (I–60 /I+60 )
from the I–V relationships (Fig. 1G, black circles) is
very similar to the rectification observed before the
current decline onset (Fig. 1G, green circles). Second,
inactivation and rectification appear to have a different pH
rectification develops almost instantaneously (Fig. 2A),
inactivation occurs within several seconds (Fig. 1D, E),
and thus within the 20 ms voltage steps used to elicit
the I–V relationship there would not be enough time
for a significant inactivation to develop. Indeed, the
A
B
pHо 5
C
pHо 7.4
pHо 5
Current (pA)
2
1
0
–1
–200 –100 0
2 nA
σ2(pA2)
270
Voltage (mV)
2 nA
+50 mV
180
90 i = 0.36
0
0
90
100
I (pA)
–50 mV
0
+100 mV
+150 mV
+180 mV
1800
1800
1200
1200
900 i = 0.88
900
600
600
300
i = 1.73
i = 2.19
0
0
0
0 250 500 750
0
500 1000
0
500 1000
I (pA)
+100 mV
0
0
–100 –200
I (pA)
pHо 5
–25 –50
I (pA)
pHо 7.4
C–
C–
Ob –
O–
1s
0.2 s
H
i = 2.1 pA
H+i
δ
H+
-BS
(1-δ)
reaction coordinates
Punblocked/Pmax
U–2
K(0)–1
U1
H+о
1
0
–1
–2
–3
–4
–5
i = –0.6 pA
i = –2.3 pA
1.0
K(0)1
energy
i = 3.2 pA
I
K(0)2
K(0)–2
200 pA
–25 –50
I (pA)
pHо 7.4
–100 mV
pHо 5
0
5
4
3
2
1
0
–1
O–
pHо 5
45 i = –0.72
0
0
Ob –
F
–180 mV
90
i = –0.5
45
0
I (pA)
–150 mV
90
180 i = –0.22
–25 –50
I (pA)
E
I (pA)
–100 mV
360
45 i = –0.15
0
G
100 200
5 ms
D
σ2(pA2)
Figure 2. Proton block and current rectification in WT
CNGA1 channel
A and B, macroscopic current recordings in the inactivated
state obtained 2 min after the addition of 1 mM cGMP at
0 mV elicited by voltage steps from −200 to +200 mV
(V = 20 mV) at pHo 5 (A) and 7.4 (B). Leak and
capacitative components were removed by subtracting
from the cGMP-activated current those records obtained in
response to the same voltage protocol but without cGMP.
C, single-channel i–V (black circles) and macroscopic I–V
relationships (red circles) obtained from noise analysis (D)
and from the recording in A, respectively. The I–V
relationship was scaled to the i flowing at +180 mV. D,
stationary fluctuation analysis at the indicated positive
(upper panels) and negative (lower panels) voltages. A
linear regression fit throughout each dataset provided the
indicated unitary current i. Currents were evoked by 30 μM
intracellular cGMP and leak currents and variance observed
in the absence of cGMP subtracted. Each point indicates
the mean current (I) and the variance of the corresponding
mean current (σ 2 ) measured from one independent
excised membrane patch. Recordings for each patch were
of duration between 0.5 and 2 s. E, single-channel current
recordings at +100 mV at the indicated pHo . Amplitude
histogram from recordings at pHo 5 is shown at the right
(grey area). Dashed brown and blue lines represent a
two-component Gaussian fit to histograms obtained at
pHo 5 and 7.4, respectively. Black dashed lines indicate the
0 current level. C, Ob and O refer to the closed, blocked
and open states respectively; i indicates the unitary current.
F, as in E but at −100 mV. G, macroscopic current
recording in the inactivated state elicited by a voltage
prepulse held at +200 mV followed by test potentials
ranging from −200 to +200 mV (V = 20 mV). No tail
currents were observed at negative membrane potentials,
suggesting a fast mechanism of proton block. Black dashed
line indicates the 0 current level. H, a sketch of the energy
landscape for a symmetrical two-barrier Woodhull model
of block. U±i , k(0)±i and δ indicate the heights of energy
barriers, rate constants at 0 mV and the electrical distance,
respectively. I, dependence of the fraction of unblocked
channels (see Methods) on V at different pHo (filled circles,
open circles, filled triangles, open triangles, open squares
and filled squares refer to pHo 9, 8, 7, 6, 5 and 4,
respectively). The red lines represent simultaneous fits of all
the data with the model shown in F with k(0)1 , k(0)–1 ,
k(0)2 , k(0)–2 and δ equal to 7.3 × 107 M−1 s−1 , 40.8 s−1 ,
8.9 s−1 , 1.6 × 107 M−1 s−1 and 0.41, respectively.
i
l norm
0.8
0.6
0.4
0.2
0.0
–200 –100
0
100 200
Voltage (mV)
2 ms
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
862
A. Marchesi and others
this mechanism, we estimated the single-channel current
i by noise analysis of macroscopic currents at different
voltages (Fig. 2D). Estimates of the corresponding i thus
obtained are plotted as a function of voltage in Fig. 2C
(black dots). As with the macroscopic I–V relationship
(Fig. 2C, red dots), the i–V plot is outwardly rectifying
and the two curves are almost overlapping. Indeed, the
ratio of the macroscopic and unitary currents flowing at
+180 and −180 mV (I+180 /I–180 and i+180 /i–180 ) were 4.5
and 3, respectively, suggesting that the voltage dependency
of proton blockage is responsible for a large fraction of the
observed rectification.
Estimates of single-channel currents based on
stationary fluctuation analysis could be biased by filtering
settings (Hille, 1992), and therefore we also attempted to
measure single-channel events from membrane patches
containing possibly only one channel. Such electrical
recordings are shown in Fig. 2E and F and have been
obtained at +100 (Fig. 2E) and −100 mV (Fig. 2F) at
the usual pHo of 7.4 (right traces) and when pHo was
lowered to 5 (left traces). Proton elevation resulted in a
reduction of the single-channel amplitude that was more
prominent at negative voltages (Fig. 2F). At −100 mV,
amplitude histograms indicate that the i reduced from
dependency: while a significant increase in rectification
was already observed when pHo is lowered from 8
to 7 (Fig. 1B, C, G), an appreciable inactivation was
not detected until pHo was lowered to 6 (Fig. 1B–D).
Lastly, the voltage dependency of inactivation adds to the
instantaneous rectification resulting in an enhancement of
the steady-state current rectification (Fig. 1H). Thus, the
voltage- and time-dependent loss of conductance is not at
the origin of the outward rectification observed in the I–V
relationship, and the two processes of rectification and
inactivation could have different underlying molecular
mechanisms.
Outward rectification arises from voltage
dependency of proton blockage
The almost instantaneous rectification at pHo 5 (Fig. 2A)
was substantially relieved at the usual pHo 7.4 (Fig. 2B) and
could reflect obstruction of the permeation pathway by
protons within the transmembrane electrical field. Indeed,
at hyperpolarized membrane potentials, protons are
expected to be driven into the channel pore, enhancing the
block and resulting in an outward rectification. To confirm
A
B
pHо 5
J Physiol 593.4
15
*
12
τ (s)
Na+
Rb+
Cs+
9
6
3
0
C
NH4+
**
0.6
0.4
0.2
0.0
4 +
N
a+
R
b+
C
s+
M
A+
N
H
5s
(NH4+)o
(NH4+)o
i
i
a+
4 +
)i
H
N
Iss/Ipeak
N
H
4 +
)i
10 s
**
0.8
0.6
0.4
0.2
0.0
i
20 pA
**
15
12
9
6
3
0
a+
τ (s)
pHo 5
Na+
(N
(NH4+)i
(N
D
**
0.8
Iss/Ipeak
MA+
Figure 3. Properties of inactivation under
different ionic conditions in WT CNGA1
channel
A, current recordings during a voltage step at
−60 mV at pHo 5 in symmetrical 110 mM Na+
(green trace), Rb+ (brown trace), Cs+ (purple
trace), MA+ (red trace) and NH4 + (blue trace)
solutions. Currents were evoked by 1 mM cGMP
and scaled for comparison. B and C,
inactivation time constant τ (B) and residual
fractional current Iss /Ipeak (C) in the presence of
Na+ , Rb+ , Cs+ , MA+ and NH4 + . ANOVA and
Holm–Sidak post hoc tests were performed to
compare each ion against each other
(∗ P < 0.05, ∗∗ P < 0.01). D, current recordings
during a prolonged voltage step at −60 mV at
pHo 5 in symmetrical NH4 + (left trace) and
when intracellular NH4 + was replaced by an
equimolar amount of Na+ (right trace). A 2 min
wash-out between the two recordings was
performed to ensure complete recovery from
inactivation. Currents were evoked by 1 mM
cGMP. Inactivation time constant τ and residual
fractional current Iss /Ipeak in the presence of
symmetrical NH4 + and when intracellular NH4 +
was replaced by an equimolar amount of Na+
are shown in the right insets. An unpaired
two-tailed t-test was performed to compare
Na+ i to (NH4 + )i (∗∗ P < 0.01). Dashed lines in A
and D indicate the 0 current level.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
Proton transfer unlocks inactivation in CNGA1 channels
−2.3 to −0.58 pA when pHo was lowered from 7.4 to 5
(Fig. 2F), and a substantial relief of block was observed
at +100 mV (Fig. 2E, left trace). Indeed, the ratio of
the single-channel conductance at +100 and −100 mV
A
863
(γ +100 /γ –100 ) is equal to 3.6 at pHo 5. These data suggest
that a fast mechanism of proton blockage is the leading
cause of the current rectification, a notion that is further
substantiated by the lack of tail currents when classical
B
WT
C
E363A
4
τ (s)
J Physiol 593.4
pHо 5.0
2
pHо 5.0
0
D
Iss/Ipeak
0.
0.
04
08
pHо 7.4
pHо 7.4
0.
00
5s
E
pHо 7.4
7.4 pHо
0 mV
–50 mV
It/Imax
1.0
E363A
0.5
0.0
20 pA
F
20 s
G
τWT= 22.1 s
τE363A = 23.5 s
0
50 100
Time(s)
**
τ (s)
20
E363A, pHо 5.0
WT, pHо 5.0
H
10
**
n.d.
0
1.5
*
Iss/Ipeak
**
T360A, pHо 5.0
T359A, pHо 5.0
1.0
0.5
**
W
E3 T
63
T3 A
59
A
T3
60
A
0.0
2s
J
T359A pHо 5
1.0
WT
T359A
G/G+200
I
0.5
0.0
–200
50 pA
Figure 4. Properties of inactivation in pore
mutant channels
A and B, current recordings during a voltage step at
−60 mV at pHo 5 and 7.4, in symmetrical 110 mM
Na+ for the WT (A) and for the E363A mutant (B)
channels. Currents were evoked by 1 mM cGMP and
scaled for comparison. C and D, inactivation time
constant τ (C) and residual fractional current Iss /Ipeak
(D) for the E363A mutant channel at pHo 5 and 7.4.
An unpaired two-tailed t-test was performed to
compare the two conditions. No statistically significant
differences were found. E, current recording during a
voltage step at −50 mV at pHo 7.4 in symmetrical
Na+ for the E363A mutant channel. Currents were
evoked by the application of 1 mM cGMP as indicated
by solid black bars. The inset shows the fractional
recovery It /Imax for the E363A mutant channel at pHo
7.4 (black circles) and WT channel at pHo 5 (green
circles) plotted as a function of the cumulative
interpulse interval. Data were fitted to a single
exponential function providing the indicated time
constants τ . F, current recordings during a voltage
step at −60 mV in symmetrical 110 mM Na+ for
E363A (black trace), WT (green trace), T360A (blue
trace) and T359A (red trace) channels, at pHo 5.
Currents were evoked by 1 mM cGMP and scaled for
comparison. G and H, inactivation time constant τ (G)
and residual fractional current Iss /Ipeak (H) for WT,
E363A, T359A and T360A channels. ANOVA and
Holm–Sidak post hoc tests were performed to
compare each ion against each other (∗ P < 0.05;
∗∗ P < 0.01). I, macroscopic current recording in
symmetrical 110 mM Na+ , evoked by 1 mM cGMP and
elicited by voltage steps from −200 to +200 mV
(V = 20 mV) at pHo 5 for the T359A mutant
channel. J, dependence of G/G+200 on V for WT
(black dots) and T359A mutant (red squares) channels
at pHo 5. While abolishing desensitization (F), Thr359
replacement with an alanine does not affect voltage
dependency of proton blockage (J). Dashed black lines
in A, B, E and F indicate the 0 current level.
5.0
5 ms
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
–100
0
100
Voltage (mV)
200
864
A. Marchesi and others
voltage jump experiments are considered to uncover the
voltage dependency of channel deactivation (Fig. 2G).
Like divalent cations, protons appear to reduce unitary
currents by acting inside the conduction pathway of the
channel (Dzeja et al. 1999; Seifert et al. 1999), and therefore
we examined whether the Woodhull model (Woodhull,
1973) describes proton blockage (Fig. 2H). We assumed
that H+ binds to a binding site (BS) located at a certain
electrical distance δ from the extracellular side of the
membrane (Fig. 2H). Figure 2I illustrates the normalized
G/G+200 relationship recorded at different pHo , the shape
of which could be described by the Woodhull model,
yielding a δ of 0.41. In this framework, the negative
slope of the conductance observed at negative voltages
(Fig. 2I) reflects the punch-through of H+ from the pore.
The model, however, does not satisfactorily fit the data at
a pHo equal to or lower than 5 (filled and open squares,
respectively, in Fig. 2I), in which the predicted proton
A
+10 mV
J Physiol 593.4
blockage is consistently larger than effectively measured.
A very similar model has been described to explain
extracellular divalent cation blockage and the blocking
site is located at almost the same electrical distance δ of
0.45 (Seifert et al. 1999). It is thus conceivable that protons
and divalent cations compete for the same BS within the
selectivity filter of CNG channels.
The inactivation gate is located at the selectivity filter
In K+ channels, the intimate relationship between the
so-called C-type inactivation and the selectivity filter
is illustrated by the substantial effects of the permeant
ions on inactivation gating (López-Barneo et al. 1993;
Starkus et al. 1997). Therefore, we examined whether
ionic substitution could also affect the extent and
the rate of inactivation in CNGA1 channels. Current
decline was observed in the presence of Na+ , NH4 +
+30 mV
+50 mV
+70 mV
−30 mV
−50 mV
−70 mV
0.5 nA
5s
−10 mV
B
C
It/Imax
1.0
D
I/I+90
+70 mV
1.0
1.0
+30 0.5
0.5
−30 0.0
−70 mV
0.0
0
5 10 15
Time (s)
−0.5
1−Iss/Ipeak
−60 mV
+60 mV
0.5
Ipeak
Iss
−100 −50 0 50 100
Voltage (mV)
0.0
5
6
7
8
pHo
Figure 5. Voltage dependency of inactivation in WT CNGA1 channel
A, current recordings at pHo 5 from the same inside-out patch at the indicated membrane potentials in symmetrical
Na+ . Currents were evoked by 1 mM cGMP. Dashed line indicates the 0 current level. B, time course of inactivation
It /Imax at pHo 5 for the indicated membrane potentials from experiments as in A where sampling points were
averaged every 500 μs. Each current decay was obtained from at least three independent experiments. Imax was
measured at the peak of the cGMP current. C, dependence of the cGMP-activated current on membrane voltage
prior to current decline (black circles) and after completion of inactivation (white circles). Current was scaled to
the current flowing at +90 mV. D, dependence of fractional inactivation (1 – Iss )/Ipeak at + 60 mV (white dots) and
−60 mV (black dots) on pHo . Ipeak and Iss were measured at the beginning and termination of the cGMP pulse,
respectively. Blue, green and red lines refer to simultaneous fits of all the data with schemes i, ii, and iii shown in
Fig. 6A, respectively. For scheme i association constant B(0) was 1.59 × 104 M−1 (pKa 4.2). The electrical distance
δ B was 0.37. The forward (i+ ) and backward (i– ) inactivation rate constants were 0.41 and 0.05 s−1 , respectively.
For scheme ii association constants B(0)1 and B(0)2 were 2.8 × 105 M−1 (pKa 5.44) and 2.5 × 104 M−1 (pKa 4.4),
respectively. The electrical distance δ B was 0.22. The forward (i+ ) and backward (i– ) inactivation rate constants were
0.51 and 0.035 s−1 , respectively. For scheme iii association constants P(0), B(0)1 and B(0)2 were 1.59 × 104 M−1
(pKa 4.2), 1.25 × 106 M−1 (pKa 6.1) and 5 × 104 M−1 (pKa 4.7), respectively. The electrical distances δ P and δ Bi
were 0.37 and 0.45, respectively. The forward (i+ ) and backward (i– ) inactivation rate constants were 0.41 and
0.05 s−1 , respectively.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
J Physiol 593.4
Proton transfer unlocks inactivation in CNGA1 channels
and methylammonium (MA+ ) (Fig. 3A), although
inactivation was appreciably slower in the presence of
NH4 + (τ = 10.5 ± 1.7 s, n = 5) compared to Na+
(τ = 7.5 ± 1.2 s, n = 5) and MA+ (τ = 7.2 ± 1.2 s, n = 3)
conditions (Fig. 3B). Moreover, in the presence of either
NH4 + or MA+ ions, current decline was significantly
less complete (Fig. 3C). When Na+ ions were replaced
with Rb+ and Cs+ , inactivation properties did not differ
among the different conditions (Fig. 3A–C). Thus, the
pore occupancy of specific ions appears to be one of the
microscopic factors controlling inactivation. Recently we
showed that the presence of large cations such as Rb+
and Cs+ in the bathing medium inhibits the Na+ inward
current by interacting with Thr359 and Thr360 at the
intracellular mouth of the selectivity filter. Indeed, the
ring of these threonines contributes to an ion binding
site at the intracellular entrance of the selectivity filter,
and cations present in the intracellular medium could
potentially interact with this site and affect channel gating
and permeation (Marchesi et al. 2012). Therefore, we
investigated whether occupancy of this binding site by
certain ions compared to others could affect inactivation
properties. When NH4 + ions in the bathing medium were
replaced by an equimolar amount of Na+ , the NH4 +
current not only appeared to be inhibited by the presence of intracellular Na+ as the NH4 + peak current
reduced by 32% in the presence of intracellular Na+
(I(NH4 + )i /I(Na+ )i = 0.68 ± 0.06; n = 4), but inactivation
also proceeded faster (τ = 5.38 ± 0.98 s, n = 4) and
was more complete (Fig. 3D). It is therefore conceivable
that Na+ occupancy of this intracellular site, signalled by
the NH4 + current blockage, prompted channel collapse
towards the non-conductive, inactivated state.
Previous studies identified the Glu363 residue at the
outer mouth of the selectivity filter as the major proton
and divalent cation target (Root & MacKinnon, 1993,
1994; Eismann et al. 1994; Rho & Park, 2013; Morrill
& MacKinnon, 1999). Therefore, we evaluated whether
besides current rectification, proton binding to Glu363
also controls the time- and voltage-dependent loss of
conductance discussed earlier. Similar to WT channels
when pHo was reduced to 5, mutant E363A channels
inactivated at the usual pHo of 7.4 (Fig. 4A, B, E), as
previously described (Mazzolini et al. 2009). Moreover,
the current decline observed at pHo 7.4 and pHo 5 is
almost identical in this mutant (Fig. 4B); the inactivation
time constant τ is equal to 3.1 ± 0.1 and 2.9 ± 0.2 s,
respectively, and the residual steady-state current to
0.04 ± 0.01 and 0.05 ± 0.01 (Fig. 4C, D; n 4). Thus,
neutralization of Glu363 abolishes the pH dependency of
inactivation, indicating that proton binding to Glu363 is
also responsible for the slow current decline. Remarkably,
compared to the WT channel, the onset rate of current
decline was faster (Fig. 4F, G) and inactivation was more
pronounced in the E363A mutant channel (Fig. 4F, H),
865
while the time course of recovery did not significantly
differ in the two channels (Fig. 4E, inset). Inactivation
kinetics is from two to three orders of magnitude slower
than the gating transitions: indeed CNGA1 channels open
in a few milliseconds in response to cGMP concentration
jumps (Nache et al. 2006). Relying on this separation of
timescales, the onset rate of inactivation depends on both
the forward and the backward inactivation rate constants,
while the recovery process depends solely on the backward rate constant. As the current decline onset in the
E363A mutant is faster but recovery from this effect is
not different from the WT channel, the mutation appears
to affect only the forward inactivation rate constant, i.e.
mutation of Glu363 destabilizes the open conformation of
the pore.
To further substantiate the notion that the occupancy
of an ion binding site in close proximity to the central
cavity controls inactivation, we studied the current decline
in Thr359 and Thr360 mutant channels (Marchesi et al.
2012). Inactivation kinetics for WT, T359A and T360A at
pHo 5 are shown in Fig. 4F. Current decline developed
significantly more slowly in the T360A mutant channel
(τ = 19.0 ± 1.9 s, n = 3, Fig. 4G) compared to the WT
channel (τ = 7.5 ± 1.2 s, n = 5, Fig. 4G). When Thr359
at the intracellular entrance of the selectivity filter was
mutated to an alanine, a significant inactivation was not
observed within 20 s (Fig. 4F, H). The lack of current
decline observed in T359A channels is unlikely to arise
from a decreased affinity to protons, as the outward
rectification at pHo 5 is very similar in WT and T359A
channels (Fig. 4I, J).
Voltage dependency of inactivation is expected from
proton binding within the electrical transmembrane
field
We studied the effect of membrane potential on
inactivation kinetics by eliciting currents in response
to 1 mM cGMP at different voltages (Fig. 5A). Two
key differences among these records are the amplitude
of steady-state currents and the rate of inactivation:
the former increases as the membrane is made more
depolarized, while the latter decreases. This finding is best
illustrated in Fig. 5B, in which current decays obtained
at different voltages were normalized to the current
measured and compared immediately after addition of
cGMP. Fitting the time course of inactivation at +70
and −70 mV with a single exponential yields a τ of
inactivation equal to 16.3 and 6.7 s, respectively, and a
residual steady-state current (It /Imax ) equal to 0.63 and
0.27. The I–V relationships measured immediately after
exposure to cGMP (filled circles) and after development of
current decline (open circles) are shown in Fig. 5C and had
a different degree of rectification. Indeed, at steady state,
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
866
A. Marchesi and others
the outward rectification increased due to the additional
contribution of the inactivation (Fig. 5C). Finally, the pH
dependency of inactivation at +60 mV (open circles) and
−60 mV (solid circles) is shown in Fig. 5D. The fraction
of channels residing in the inactivated state appears to
be larger at negative membrane potentials and low pH
(Figs 1H and 5D).
The slow time course of current decline described
here might reflect a reduced accessibility of Glu363
carboxylate to extracellular protons. Indeed, previous
A
i
Hl
i+
C
O
C
O
[H+]xB
i−
HB
ii
[H+]x2xB1
HB
iii
i−
HHB
i−
HO
+
[H+]xB2
[H ]xB1
+
HB
B
[H ]xP
Hl
i+
HHl
i+
i−
HHB
pHo 7.4
pHo 5
+cGMP
20 s
i
C
ii
+cGMP
O+HO
0.4
Normalized
occupancy
Occupancy
iii
HI+HHI
0.6
0.2
0.0
0
1.0
τ= 6.5
0.5
τ= 6.5
0.0
10
10
20
Time (s)
20
Time (s)
D
E3663
H+
H+
T359
T360
structure–function studies suggested that this residue is
not directly accessible to extracellular solvent (Sun et al.
1996; Mazzolini et al. 2009; Martı́nez-François et al. 2009).
Yet, this simple interpretation is hard to reconcile with
several experimental observations. First, according to this
idea, the disruption of the proton binding site should
have resulted in a non-inactivating phenotype at pHo
5. This is in contrast to the finding that neutralization
of Glu363 results in inactivating currents at the usual
pHo of 7.4 (Fig. 4E). Indeed, the inactivation onset
and recovery in the E363A mutant are still slow and
comparable to those observed in the WT channel (Fig. 4E,
F). Second, recovery from inactivation is fast – less than
1 s as no obvious kinetics could be resolved – when
pHo is reverted from 5 to 7.4 in outside-out patches
(Fig. 1F, right panel). If the slow current run-down
reflected the low accessibility of Glu363 to protons, then
one would have expected a similar slow recovery upon
pHo switch to 7.4. Therefore, we suggest that current
decline reflects proton-induced conformational changes,
of which the kinetics are limited by a transition downstream from protonation. In this view, protons first bind
+
[H ]xB2
[H+]xP
O
C
HHl
i+
+
[H ]x1/2xB2
J Physiol 593.4
pKa = 6.1
pKa = 4.2
Figure 6. Proposed mechanism for proton action in CNGA1
channels
A, schemes showing the proposed mechanisms for proton binding
and inactivation with one proton binding site (i), two equivalent
proton binding sites (ii) and two non-equivalent and
non-independent binding sites (iii). In iii columns depict protons
(cyan circle and symbols) binding within permeation pathway and
blocking ion conduction (D). A second proton (blue circle and
symbols) binds to E363 carboxylate, possibly impairing an
electrostatic interaction between E363 side chain and neighbouring
residues in the P-helix (D) (Mazzolini et al. 2009; Martı́nez-François
et al. 2010). The open conformation of the pore is now metastable
and collapses to inactivated states H I and HH I. C, O, B and I indicate
closed, open, blocked and inactivated states, respectively. H
subscripts indicate proton transfer to the pore. P and Bi are the
association constants for protons to the two non-equivalent binding
sites within the pore lumen. i+ and i– are the forward and backward
inactivation rate constants. B, simulated currents with schemes i
(blue trace), ii (green trace) and iii (red trace) shown in A in response
to a pHo step from 7.4 to 5 (black line) at −50 mV. The dotted black
line indicates the mean fractional peak current observed in
outside-out patches at pHo 5 (I(pHo 5)/I(pHo 7.4) = 0.21 ± 0.02,
n = 3; Fig. 1F). The dashed black line indicates the 0 current level.
Rate constants are listed in the legend to Fig. 5. C, time course of
occupancies for the conductive states (O + H O; solid red line) and
inactivated states (H I + HH I; solid brown line). The occupancies were
calculated from the scheme iii shown in A at −50 mV and pHo 5
after a step in cGMP concentration from 0 to 1 mM. The inset at the
right shows the decaying and rising phase of these occupancies,
highlighting the same time constants (τ = 6.5 s). cGMP application
in B and C is indicated by the solid grey lines. D, cartoon illustrating
CNGA1 pore domain. The inset shows an enlargement of the
selectivity filter region. Residues E363, T359 and T360 are shown.
Cyan and blue circles represent two protons binding, respectively,
within the permeation pathway and to the E363 side chain. The
logarithmic acidic dissociation constants (pKa ) for the two binding
sites were determined from P(0) and B1 (0) association constants by
fitting the model in A to the data shown in Fig. 5.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
J Physiol 593.4
Proton transfer unlocks inactivation in CNGA1 channels
to Glu363 within the electrical transmembrane field,
quickly blocking ion conduction and destabilizing the
pore, which subsequently undergoes a slow structural
collapse leading to a virtually closed channel. This
molecular picture is captured by the simple linear scheme
shown in Fig. 6Ai, involving transitions among an open,
blocked and inactivated state, which could successfully
describe the pH and voltage dependency of inactivation,
yielding a pKa of 4.2 (Fig. 5B, D blue lines). Although
we could not rule out the possibility that membrane
voltage might affect the conformation of charged residues
and/or structures located within the electrical field as
suggested for bacterial K+ channel KcsA (Cordero-Morales
et al. 2006), the properties of the observed current
decline are well accounted by the voltage-dependent
concentration of H+ within the electrical field alone,
without the necessity of taking explicitly into account a
voltage-dependent conformational change. The fraction
of the membrane voltage at the binding site thus obtained
(δ 0.37) is consistent with the expected location of
Glu363 within the electrical field (Contreras et al. 2010).
This scheme, however, appears to be critically deficient
in reproducing the instantaneous and steady-state I–V
relationship (Figs 5C and 6B, blue lines). The basis
of this shortcoming is the different pH dependency
of proton blockage and inactivation. Indeed, proton
blockage has been associated with an apparent acidic
dissociation constant (pKa ) between 6 and 7 in several
reports (Root & MacKinnon, 1994; Seifert et al. 1999). It is
therefore possible that the binding of at least two protons
underlines proton blockage and inactivation in CNGA1
channels.
Discussion
Desensitization and inactivation are widespread
phenomena in ion channels, in which they modulate the
amplitude, duration and frequency of electrical signalling
within and between cells. Unlike many of their cousin K+
channels and other membrane receptors, CNG channels
do not desensitize or inactivate at physiological pH when
a network of chemical interactions between residues in
the pore region is maintained (Mazzolini et al. 2009). We
have shown that in response to changes of extracellular
pH, CNG channels also inactivate, representing a
possible novel regulatory mechanism that has so far been
neglected.
Previous studies suggested the presence of multiple
H+ -binding sites in the CNG channel pore. Single-channel
current analysis in olfactory CNG channels indicated that
the binding of two protons is required to model the proton
monovalent current inhibition (Root & MacKinnon,
1994). Moreover, Seifert et al. (1999, fig. 5B, C) showed that
a Hill coefficient of 2 describes the pH dependency of
divalent cation blockage in CNGA1 channels, suggesting
867
the binding of at least two protons. In agreement with
these earliest observations we were unable to describe
the experimental data when the binding of only one
proton was considered (Fig. 5). Similarly, a linear scheme
assuming two equivalent proton binding sites (Fig. 6Aii)
failed to satisfactory reproduce the experimental data as
it overestimates the current inhibition observed at low
pHo (compare Figs 1F and 6B, green trace). Therefore,
we evaluated a more general kinetic scheme in which
two non-equivalent and non-independent proton binding
reactions were modelled (Fig. 6Aiii). This model appears
to adequately describe key features of the reported current
decline, such as the time course of inactivation (Fig. 5B,
red lines), its pH dependency (Fig. 5D, red lines), the
instantaneous and steady-state rectification (Fig. 5C, red
lines) and the fractional current blockage observed at
pHo 5 (Fig. 6B, red trace). As we were unable to resolve
the kinetics of H+ unbinding in outside-out patches
(Fig. 1F) as well as the kinetics of proton blockage from
voltage jump experiments (Fig. 2G), the proton binding
reactions are not likely to contribute to the observed slow
current relaxation kinetics and the two binding reactions
are therefore assumed to be diffusion limited in this
model. Once protons have bound, the slow current decline
reflects the very slow inactivation rate constants, which
are the rate-limiting reactions in the scheme. Indeed,
the time courses of the occupancies for the conductive
states (O + H O, red lines) and the inactivated states
(H I + HH I, brown lines) are symmetrical (Fig. 6C),
yielding at −50 mV the same time constant τ of 6.5 s.
Thus, we propose that the non-equivalent binding of two
protons within the transmembrane electrical field underlies voltage-dependent blockage and inactivation.
It has also been suggested that voltage-dependent gating
contributes to the rectification of the macroscopic currents
at very low pHo (pHo = 4) as a consequence of Glu363
side chain titration (Martı́nez-François et al. 2010). Based
on noise analysis (Fig. 2C, D), single-channel recordings
(Fig. 2E, F) and voltage-jump experiments (Fig. 2G), this
mechanism does not appear to significantly contribute
to the outward rectification observed at higher pHo
(pHo 5) and has therefore not been included (Fig. 6A).
How could the proposed kinetic mechanism be interpreted in the light of the available structural and functional
data? The crystal structure of a recently solved bacterial
channel mimicking the CNG channel pore (Derebe et al.
2011) suggests that Ca2+ ions are chelated by backbone
carbonyl oxygens and not by the Glu363 side chain, which
is engaged in hydrogen bonding interactions with its
neighbouring residues and buried underneath the external
surface of the protein. The authors of this study suggested
that the Glu363 carboxylate negative charge could perturb
the electron shell distribution along the backbone of the
filter residues, making certain carbonyl oxygen atoms,
such as those of Gly65, more electronegative and suited
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
868
A. Marchesi and others
for Ca2+ binding (Derebe et al. 2011). These observations
are consistent with several recent structure–function
studies, suggesting that interactions between the Glu363
side chain and residues in the P-helix are necessary for
normal gating in CNG channels (Mazzolini et al. 2009;
Martı́nez-François et al. 2009, 2010). In this view proton
blockage could arise from the high-affinity binding to the
main chain carbonyl oxygens within the selectivity filter
(Fig. 6D, pKa = 6.1). A second, low-affinity binding site
for protons could be constituted by the Glu363 side chain
(pKa = 4.2), titration of which results in the neutralization
of the negatively charged carboxylate and in the weakening
of key interactions anchoring the selectivity filter to
the surrounding channel moiety (Fig. 6D) (Mazzolini
et al. 2009; Martı́nez-François et al. 2009, 2010). After
Glu363 protonation, glycine carbonyl oxygen specificity
for protons and divalent cations is lost – the pKa of the
proton high-affinity binding site reduces from 6.1 to 4.7
– and thus a substantial relief from blockage is observed
at low extracellular pHo (Fig. 2I). It then follows a slow
pore collapse toward a non-conductive, inactivated state.
These observations suggest that the two binding sites
are non-independent, and explain why neutralization of
the same residue (Glu363) affects both proton blockage
(Root & MacKinnon, 1994) and the pH dependency of
inactivation (Fig. 4).
One of the hallmarks of the C-type inactivation is
its intimate dependence on the permeant ion. Indeed,
it has long been known that the rate and extent of
C-type inactivation is governed by the occupancy of
an ion binding site near the external mouth of the
pore via a foot-in-the-door mechanism (Yellen, 1998;
Kurata & Fedida, 2006). Our results indicate that such
a relationship between the permeant ion and inactivation
also exists in CNG channels (Fig. 3), although the underlying mechanism appears to be somewhat different. In
fact, inactivation appears to be primarily modulated by
the nature of the intracellular cation, being faster in the
presence of intracellular Na+ (Fig. 3D). Furthermore,
inactivation is abolished when the intracellular ion
binding site is disrupted (Fig. 4F), thus suggesting that
the occupancy of an ion binding site possibly equivalent
to the fourth ion binding site within the selectivity
filter (S4 site) of the K+ channel might control the rate
and extent of inactivation in CNG channels. Therefore,
it is conceivable that the pore collapse triggered by
the protonation of Glu363 implies fine conformational
changes also of the intracellular vestibule involving
the two threonines Thr359 and Thr360. Although the
molecular basis underpinning the C-type inactivation
is still of debate (Devaraneni et al. 2013; Armstrong
& Hoshi, 2014), according to the predominant view
inactivation is associated with a conformational change
close to the external mouth of the K+ channel pore
J Physiol 593.4
(Yellen, 1998; Kurata & Fedida, 2006; Hoshi & Armstrong,
2013). Nonetheless, rearrangements near the intracellular
face of the selectivity filter have been implied as well.
Indeed, Posson et al. (2013) demonstrated that the
binding affinity of certain intracellular blockers depends
on the conformational state of the selectivity filter
(open-conductive versus closed-inactivated). The authors
of this study argued that the observed state-dependent
affinities reflect a conformational change at the intracellular face of the selectivity filter upon inactivation.
These rearrangements close to the intracellular end of
the selectivity filer could be more pronounced during the
CNG channel inactivation process as they may reflect the
allosteric connection between the CN-binding domain
and the selectivity filter gate. In this view, residues
Thr359 and Thr360 might play an important role in the
observed current decline. Overall, inactivation appears to
be associated with a local remodelling confined to the
selectivity filter and neighbouring regions in CNG as well
as K+ channels (Roncaglia & Becchetti, 2001; Kurata &
Fedida, 2006; Mazzolini et al. 2009).
In the present study, we demonstrate that the outward
rectification observed at pHo 5 at stationary conditions
(i.e. after a steady exposure to cGMP for 1–2 min) is
mainly caused by asymmetries in the unitary current
(Fig. 2). This finding is consistent with several earlier
studies suggesting that proton elevation is associated with
the stabilization of low conductance states primarily at
negative voltages (Root & MacKinnon, 1994; Rho &
Park, 2013; Morrill & MacKinnon, 1999). We propose
that a Woodhull model of block, alongside divalent
cation blockage (Seifert et al. 1999), underlies proton
block in CNG channels. This is directly opposite to the
conclusions reached by a recent report suggesting that
the outward rectification is due to an inherent channel
voltage-dependent gating (Martı́nez-François et al. 2010).
The reasons for these discrepancies are not clear.
Several of the Glu363 mutants display voltage-dependent
gating in addition to inactivation (Bucossi et al. 1996;
Martı́nez-François et al. 2009). It is thus likely that voltage
gating additively contributes to the observed rectification
when extracellular pH is further lowered to 4 and slow
activation kinetics are observed (Martı́nez-François et al.
2010). Determining the extent that such a mechanism
contributes to the outward rectification at very low
pHo will require further experimentation. Nonetheless,
demonstration of common gating features (inactivation
and voltage gating) as the result of pore mutation and
protonation further illustrates the gating role of the outer
pore in CNG channels.
Changes in pHo can arise in a variety of physiological and pathophysiological conditions, such as neuronal activity, ischaemia and inflammation (Kellum et al.
2004; Isaev et al. 2008; Magnotta et al. 2012). Low pH
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
J Physiol 593.4
Proton transfer unlocks inactivation in CNGA1 channels
acts as a negative feedback mechanism that inhibits the
CNGA1 channel in a state-dependent manner and may
represent an unrecognized endogenous signal regulating
CNG physiological functions in diverse tissues.
References
Armstrong CM & Hoshi T (2014). K+ channel gating: C-type
inactivation is enhanced by calcium or lanthanum outside. J
Gen Physiol 144, 221–230.
Becchetti A, Gamel K & Torre V (1999). Cyclic nucleotide-gated
channels. Pore topology studied through the accessibility of
reporter cysteines. J Gen Physiol 114, 377–392.
Bucossi G, Eismann E, Sesti F, Nizzari M, Seri M, Kaupp UB &
Torre V (1996). Time-dependent current decline in cyclic
GMP-gated bovine channels caused by point mutations in
the pore region expressed in Xenopus oocytes. J Physiol 493,
409–418.
Bucossi G, Nizzari M & Torre V (1997). Single-channel
properties of ionic channels gated by cyclic nucleotides.
Biophys J 72, 1165–1181.
Contreras JE, Chen J, Lau AY, Jogini V, Roux B & Holmgren M
(2010). Voltage profile along the permeation pathway of an
open channel. Biophys J 99, 2863–2869.
Cordero-Morales JF, Cuello LG & Perozo E (2006).
Voltage-dependent gating at the KcsA selectivity filter. Nat
Struct Mol Biol 13, 319–322.
Craven KB & Zagotta WN (2006). CNG and HCN channels:
two peas, one pod. Annu Rev Physiol 68, 375–401.
Derebe MG, Zeng W, Li Y, Alam A & Jiang Y (2011). Structural
studies of ion permeation and Ca2+ blockage of a bacterial
channel mimicking the cyclic nucleotide-gated channel pore.
Proc Natl Acad Sci USA 108, 592–597.
Devaraneni PK, Komarov AG, Costantino CA, Devereaux JJ,
Matulef K & Valiyaveetil FI (2013). Semisynthetic K+
channels show that the constricted conformation of the
selectivity filter is not the C-type inactivated state. Proc Natl
Acad Sci USA 110, 15698–15703.
Dmitriev AV & Mangel SC (2001). Circadian clock regulation
of pH in the rabbit retina. J Neurosci 21, 2897–2902.
Dzeja C, Hagen V, Kaupp UB & Frings S (1999). Ca2+
permeation in cyclic nucleotide-gated channels. EMBO J 18,
131–144.
Eismann E, Müller F, Heinemann SH & Kaupp UB (1994). A
single negative charge within the pore region of a
cGMP-gated channel controls rectification, Ca2+ blockage,
and ionic selectivity. Proc Natl Acad Sci USA 91,
1109–1113.
Hamill OP, Marty A, Neher E, Sakmann B & Sigworth FJ
(1981). Improved patch-clamp techniques for
high-resolution current recording from cells and cell-free
membrane patches. Pflügers Arch 391, 85–100.
Hille B (1992). Ionic Channels of Excitable Membranes, 2 Sub.
Sinauer Associates Inc., Sunderland, MA.
Hoshi T & Armstrong CM (2013). C-type inactivation of
voltage-gated K+ channels: pore constriction or dilation? J
Gen Physiol 141, 151–160.
869
Isaev NK, Stelmashook EV, Plotnikov EY, Khryapenkova TG,
Lozier ER, Doludin YV, Silachev DN & Zorov DB (2008).
Role of acidosis, NMDA receptors, and acid-sensitive ion
channel 1a (ASIC1a) in neuronal death induced by ischemia.
Biochemistry (Mosc) 73, 1171–1175.
Kaupp UB, Niidome T, Tanabe T, Terada S, Bönigk W,
Stühmer W, Cook NJ, Kangawa K, Matsuo H & Hirose T
(1989). Primary structure and functional expression from
complementary DNA of the rod photoreceptor cyclic
GMP-gated channel. Nature 342, 762–766.
Kaupp UB & Seifert R (2002). Cyclic nucleotide-gated ion
channels. Physiol Rev 82, 769–824.
Kellum JA, Song M & Li J (2004). Science review: extracellular
acidosis and the immune response: clinical and physiologic
implications. Crit Care 8, 331–336.
Kurata HT & Fedida D (2006). A structural interpretation of
voltage-gated potassium channel inactivation. Prog Biophys
Mol Biol 92, 185–208.
Leung YK, Du J, Huang Y & Yao X (2010). Cyclic
nucleotide-gated channels contribute to thromboxane
A2-induced contraction of rat small mesenteric arteries.
PLoS ONE 5, e11098.
Liu J & Siegelbaum SA (2000). Change of pore helix
conformational state upon opening of cyclic
nucleotide-gated channels. Neuron 28, 899–909.
López-Barneo J, Hoshi T, Heinemann SH & Aldrich RW
(1993). Effects of external cations and mutations in the pore
region on C-type inactivation of Shaker potassium channels.
Receptors Channels 1, 61–71.
Lopez-Jimenez ME, González JC, Lizasoain I, Sánchez-Prieto J,
Hernández-Guijo JM & Torres M (2012). Functional
cGMP-gated channels in cerebellar granule cells. J Cell
Physiol 227, 2252–2263.
Magnotta VA, Heo H-Y, Dlouhy BJ, Dahdaleh NS, Follmer RL,
Thedens DR, Welsh MJ & Wemmie JA (2012). Detecting
activity-evoked pH changes in human brain. Proc Natl Acad
Sci USA 109, 8270–8273.
Marchesi A, Mazzolini M & Torre V (2012). A ring of
threonines in the inner vestibule of the pore of CNGA1
channels constitutes a binding site for permeating ions. J
Physiol 590, 5075–5090.
Martı́nez-François JR, Xu Y & Lu Z (2009). Mutations reveal
voltage gating of CNGA1 channels in saturating cGMP. J
Gen Physiol 134, 151–164.
Martı́nez-François JR, Xu Y & Lu Z (2010). Extracellular
protons titrate voltage gating of a ligand-gated ion channel. J
Gen Physiol 136, 179–187.
Mazzolini M, Anselmi C & Torre V (2009). The analysis of
desensitizing CNGA1 channels reveals molecular
interactions essential for normal gating. J Gen Physiol 133,
375–386.
Mazzolini M, Marchesi A, Giorgetti A & Torre V (2010). Gating
in CNGA1 channels. Pflügers Arch 459, 547–555.
Morrill JA & MacKinnon R (1999). Isolation of a single
carboxyl-carboxylate proton binding site in the pore of a
cyclic nucleotide-gated channel. J Gen Physiol 114, 71–83.
Nache V, Kusch J, Hagen V & Benndorf K (2006). Gating of
cyclic nucleotide-gated (CNGA1) channels by cGMP jumps
and depolarizing voltage steps. Biophys J 90, 3146–3154.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society
870
A. Marchesi and others
Nizzari M, Sesti F, Giraudo MT, Virginio C, Cattaneo A &
Torre V (1993). Single-channel properties of cloned
cGMP-activated channels from retinal rods. Proc Biol Sci
254, 69–74.
Pavlov I, Kaila K, Kullmann DM & Miles R (2013). Cortical
inhibition, pH and cell excitability in epilepsy: what are
optimal targets for antiepileptic interventions? J Physiol 591,
765–774.
Posson DJ, McCoy JG & Nimigean CM (2013). The
voltage-dependent gate in MthK potassium channels is
located at the selectivity filter. Nat Struct Mol Biol 20,
159–166.
Rho SH & Park CS (2013). Extracellular proton alters the
divalent cation binding affinity in a cyclic nucleotide-gated
channel pore. FEBS Lett 440, 199–202.
Roncaglia P & Becchetti A (2001). Cyclic-nucleotide-gated
channels: pore topology in desensitizing E19A mutants.
Pflügers Arch 441, 772–780.
Root MJ & MacKinnon R (1993). Identification of an external
divalent cation-binding site in the pore of a cGMP-activated
channel. Neuron 11, 459–466.
Root MJ & MacKinnon R (1994). Two identical noninteracting
sites in an ion channel revealed by proton transfer. Science
265, 1852–1856.
Seifert R, Eismann E, Ludwig J, Baumann A & Kaupp UB
(1999). Molecular determinants of a Ca2+ -binding site in the
pore of cyclic nucleotide-gated channels: S5/S6 segments
control affinity of intrapore glutamates. EMBO J 18,
119–130.
Sesti F, Eismann E, Kaupp UB, Nizzari M & Torre V (1995).
The multi-ion nature of the cGMP-gated channel from
vertebrate rods. J Physiol 487, 17–36.
Starkus JG, Kuschel L, Rayner MD & Heinemann SH (1997).
Ion conduction through C-type inactivated Shaker channels.
J Gen Physiol 110, 539–550.
Sun ZP, Akabas MH, Goulding EH, Karlin A & Siegelbaum SA
(1996). Exposure of residues in the cyclic nucleotide-gated
channel pore: P region structure and function in gating.
Neuron 16, 141–149.
Tolner EA, Hochman DW, Hassinen P, Otáhal J, Gaily E,
Haglund MM, Kubová H, Schuchmann S, Vanhatalo S &
Kaila K (2011). Five percent CO2 is a potent, fast-acting
inhalation anticonvulsant. Epilepsia 52, 104–114.
J Physiol 593.4
Woodhull AM (1973). Ionic blockage of sodium channels in
nerve. J Gen Physiol 61, 687–708.
Yellen G (1998). The moving parts of voltage-gated ion
channels. Q Rev Biophys 31, 239–295.
Yu FH, Yarov-Yarovoy V, Gutman GA & Catterall WA (2005).
Overview of molecular relationships in the voltage-gated ion
channel superfamily. Pharmacol Rev 57,
387–395.
Zufall F, Shepherd GM & Barnstable CJ (1997). Cyclic
nucleotide gated channels as regulators of CNS
development and plasticity. Curr Opin Neurobiol 7,
404–412.
Additional Information
Competing interests
The authors declare that there are no competing interests.
Author contributions
M.A. and A.M. performed experiments on oocytes. M.M.
performed mutagenesis. All authors participated in designing
experiments, and analysing and interpreting the data. A.M.
wrote the paper, which was revised by all authors in
collaboration. A.M. and V.T. supervised the project. All authors
approved the final version of the manuscript.
Funding
We acknowledge the financial support of the following projects
within the Seventh Framework Programme for Research of the
European Commission: the SI-CODE project for Future and
Emerging Technologies (FET) no. FP7 - 284553, the FOCUS
Project no. FP7-ICT-270483 and the NEUROSCAFFOLDS
Project no. 604263.
C 2014 The Authors. The Journal of Physiology published by John Wiley & Sons Ltd on behalf of The Physiological Society